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In June, August, and October 2013, intact sediment cores were collected from the mudflats of the fertilized and reference creeks at low tide (n=3 per creek in June and October, n=6 per creek in August; 31 cm diameter x 15 cm deep). These time periods reflected late spring, late summer, and early fall conditions. Sediment cores were transported to Woods Hole Oceanographic Institution’s mesocosm system (Woods Hole, MA). Cores were placed in rectangular fiberglass tanks (2.7 m x 1.2 m x 0.8 m, l x w x d) that served as a water bath to minimize extreme fluctuations in day – night temperatures. The mesocosm system is located outside, so cores experienced ambient weather conditions.

Upon placement in the fiberglass tanks, the overlying water collected with each sediment core was continuously recirculated (~9 cm deep). The cores acclimated to the mesocosm system for 1 – 3 days, depending on weather, as we applied the 13C label during sunny periods when BMA would be productive. Visible epifauna (e.g., snails, shrimp) were removed to minimize grazing on benthic microbes. At the beginning of each experiment, the overlying water column was removed and replaced with filtered water from the creek where the core was collected. We used 0.2 um filtered creek water to minimize label uptake by water column microbes and recirculated the water column to maintain well-mixed conditions. The isotopic label was added as 13C-sodium bicarbonate (NaHCO3, 99 atom %, Sigma-Aldrich) to the water column of each core in June and October and half of the cores from each creek in August. The other half of the August cores received 13C-labeled S. alterniflora detritus. This material was produced from a separate experiment in which living S. alterniflora plants from PIE-LTER were dosed with 13CO2 for 3 h (Spivak & Reeve 2015). Aboveground leaves were harvested after label exposure, dried (60 deg C), and ground into a coarse powder that was evenly applied across the sediment surface. 13C-S. alterniflora was only applied in August due to availability of the labeled material. From here forward, experiments receiving 13C-NaHCO3 or 13C-S. alterniflora are referred to as BMA or S. alterniflora experiments, respectively, to reflect the autotrophic source of carbon to bacteria. On average (+/- standard error, S.E.), the cores received 11.90 +/- 0.07 mg, 10.90 +/- 0.05, and 10.97 +/- 0.09 mg 13C from NaHCO3 application in June, August, and October, respectively, and 3.17 +/- 0.12 mg 13C from the S. alterniflora detritus in August. The 13C label, as NaHCO3 or S. alterniflora, was applied for four hours between 11:00 – 15:00 h before the overlying water was removed and the cores were rinsed with at least three volumes of filtered creek water to remove unused label. The four hour sampling period was based on results from a preliminary study demonstrating that this timeframe was sufficient for detecting the label in algal and bacterial lipids. After the final rinse, the overlying water column was replaced with filtered creek water and recirculated for the duration of the experiments.

Sediment samples for organic matter composition were collected by placing a hard plastic sleeve around a polyvinyl chloride (PVC) corer (5 cm diameter x 15 cm deep) and then removing the corer. The plastic sleeve remained in place to maintain the integrity of the sediment column and mark the core location (Spivak 2015). The top 0.5 cm of each core was collected into pre-combusted vials and frozen (-80 deg C) until analysis for total organic carbon and nitrogen content and stable isotopes (d13C, d15N) and lipid biomarker composition. Adjacent samples for benthic chlorophyll were collected with smaller cores (1.5 cm diameter x 1 cm deep) into glass vials and frozen (-20 deg C) until analysis. Additional sediment cores for organic matter composition and benthic chlorophyll were collected 4, 8, 24, and 48 h after the 13C-labeled NaHCO3 was applied in June, August, and October and 4, 8, 24, and 144h after the 13C-labeled S. alterniflora was applied in August.

Lipid biomarker compounds were extracted using a modified Bligh and Dyer (1959) method. Sediment samples were extracted with a chloroform : methylene chloride : phosphate buffer saline mixture (2:1:0.8, v:v:v) using a microwave-accelerated reaction system (MARS6); samples were heated to 80 deg C for 10 min with continuous stirring. Following extraction, samples were partitioned and the organic phase was removed. The total lipid extract was concentrated under N2 and samples were separated on silica gel columns by eluting with chloroform, acetone (F1/2), and methanol (F3) (Guckert et al. 1985). The F3 (phospholipids) was dried under N2 and saponified with 0.5 M NaOH at 70 deg C for 4 h. Saponified samples were acidified and extracted three times with hexane. The extract was methylated with acidic methanol (95:5 methanol: HCl) and heated overnight at 70 deg C to form fatty acid methyl esters (FAME). Samples were analyzed with an Agilent 7890 gas chromatograph with an effluent split ~70:30 between a 5975C mass spectrometer and a flame ionization detector. Peaks were separated on an Agilent DB-5 ms column (60 m, 0.25 mm inner diameter, 0.25 um film). FAME concentrations were quantified using methyl heneicosanoate as an internal standard. FAs are designated A:BwC, where A is the number of carbon atoms, B is the number of double bonds, and C is the position of the first double bond from the aliphatic ‘w’ end of the molecule. The prefixes ‘i’ and ‘a’ refer to iso and anteiso methyl branched FAs and indicate whether the methyl group is attached to the penultimate or antepenulttimate carbon atoms (Bianchi & Canuel 2011).

A flame ionization detector (FID) is a scientific instrument that measures the concentration of organic species in a gas stream. It is frequently used as a detector in gas chromatography. Standalone FIDs can also be used in applications such as landfill gas monitoring, fugitive emissions monitoring and internal combustion engine emissions measurement in stationary or portable instruments.

Instrument separating gases, volatile substances, or substances dissolved in a volatile solvent by transporting an inert gas through a column packed with a sorbent to a detector for assay. (from SeaDataNet, BODC)

General term for instruments used to measure the mass-to-charge ratio of ions; generally used to find the composition of a sample by generating a mass spectrum representing the masses of sample components.

Capable of being performed in numerous environments, push coring is just as it sounds. Push coring is simply pushing the core barrel (often an aluminum or polycarbonate tube) into the sediment by hand. A push core is useful in that it causes very little disturbance to the more delicate upper layers of a sub-aqueous sediment.